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CORM-3 induces DNA damage through Ru(II) binding to DNA

When the ‘CO-releasing molecule-3’, CORM-3 (Ru(CO)3Cl(glycinate)), is dissolved in water it forms a range of ruthenium complexes. These are taken up by cells and bind to intracellular ligands, notably thiols such as cysteine and glutathione, where the Ru(II) reaches high intracellular concentrations. Here, we show that the Ru(II) ion also binds to DNA, at exposed guanosine N7 positions. It therefore has a similar cellular target to the anticancer drug cisplatin, but not identical, because Ru(II) shows no evidence of forming intramolecular crossbridges in the DNA. The reaction is slow, and with excess Ru, intermolecular DNA crossbridges are formed. The addition of CORM-3 to human colorectal cancer cells leads to strand breaks in the DNA, as assessed by the alkaline comet assay. DNA damage is inhibited by growth media containing amino acids, which bind to extracellular Ru and prevent its entry into cells. We conclude that the cytotoxicity of Ru(II) is different from that of platinum, making it a promising development target for cancer therapeutics.

Introduction
Organometallic complexes have long been of interest as medicinal drug targets [1]. Much of this interest centres around the platinum complex cisplatin, which (together with the related compounds carboplatin and oxaliplatin) is used in approximately half of cancer chemotherapies [1]. Cisplatin is cytotoxic because of its ability to bind covalently to DNA and crosslink it. There is however increasing incidence of resistance to cisplatin, spurring interest in other organometallics. One class of these is complexes of ruthenium.
Ruthenium exists in several oxidation states, of which the most interesting are Ru(II) and Ru(III): three Ru(III) complexes (NAMI-A, NKP-1339 and KP1019) have completed phase I clinical trials as anticancer drugs [2,3]. It is widely held that Ru(II) is the more cytotoxic oxidation state [4], and that the biological activity of Ru complexes depends heavily on the speciation of the complexes in solution: that is, on the chemical reactions undergone by the Ru ligands [2,3]. In solution, the most labile ligands tend to be halides, which dissociate rapidly to give aquated complexes [5]. Such complexes are generally found to be more soluble and more biologically active [6]. Here, we are specifically interested in Ru(II) carbonyl complexes, for which this is also true [7,8]. In blood, aquated Ru(II) complexes bind to albumin, and possibly also to the iron transport protein transferrin [9], although this has been linked more to Ru(III) than Ru(II) [2]. Binding to albumin is slow, taking many hours to reach equilibrium, and is reversible: binding to albumin has been shown to have very little effect on bioavailability, suggesting that the Ru is able to dissociate on reaching its biological target [10][11][12][13]. Albumin has therefore been described as a Ru depot [14]. Crystal structures of CORMs bound to proteins retain at least one carbonyl ligand [7,8,15], which may help in keeping the metal at a +2 oxidation level. It is still not clear how Ru complexes enter cells, though one possibility is that they use iron transport mechanisms [14]. Once inside cells, ruthenium is able to bind to proteins (via cysteine thiols [16,17] or histidines [18]), to free thiols such as glutathione [16,17], and also to DNA. In this behaviour it shows interesting differences as compared with platinum, which binds mainly to DNA, and tends to form square planar complexes, in particular linking the N7 positions of adjacent guanines, while ruthenium forms octahedral complexes [14]. The different reactivity of Ru as compared with Pt, and its greater range of biological targets, suggest that Ru may be able to evade tumour resistance better than Pt [14,19].
Ruthenium carbonyl complexes have been studied widely because of a completely different biological property: that they slowly release carbon monoxide, which itself has a wide range of biological properties. Here, we focus on one of these, carbon monoxide releasing molecule 3 [CORM-3], Ru(CO) 3 Cl(glycinate), which was designed as a rapid water-soluble carbon monoxide-releasing molecule and has been widely used in cellular studies as a source of CO [15,19,20]. For example, CORMs are being developed as drugs to deliver CO for specific therapeutic uses such as organ transplantation and treating pulmonary hypertension [21]. Numerous studies have shown that CORM-3 is toxic to bacterial cells but shows little toxicity to mammalian cells, which has stimulated research on modes of action of the Ru-based CORMs that have focussed on the toxicity of CO. However, recently we showed that CORM-3, as well as the related molecule CORM-2, in fact release very little CO in standard growth conditions [16,17,22]. Furthermore, we showed that the bacterial toxicity is due to the ruthenium(II) ion and probably not to CO at all. When CORM-3 or CORM-2 are dissolved in water, some of the Ru ligands (mainly the chloride ions as discussed above) are displaced by water or buffer, and some of the CO groups are attacked by hydroxide ions to give carboxylate ligands, producing a mixture of hydrated species [20,23]. These species are actively taken up by cells to reach high concentrations [3]. Once inside cells, they react with intracellular ligands. The most reactive ligands are thiols, such as glutathione and cysteine, which are present at high concentration inside most cells, and bind tightly (dissociation constant of the order of 1 mM) to ruthenium [16,17]. The effect of this intracellular sequestration of Ru complexes is to shift the equilibrium of hydrated Ru species from extracellular to intracellular, and produces high concentrations of Ru inside the cells, typically giving rise to intracellular Ru concentrations up to 30 to 100-fold greater than those in the medium [16,17]. One obvious consequence of the high intracellular concentration of Ru(II) is to sequester much of the free intracellular thiol, which one would expect to be a highly toxic effect, by affecting both the redox balance inside cells and glutathione-mediated detoxification mechanisms.
We have thus shown previously that binding of intracellular Ru to thiols is potentially toxic, easily sufficient to kill cells. We have however also previously shown that incubation of CORM-2 with Escherichia coli cells leads to binding of Ru to chromosomal DNA, at a level of ∼3 Ru per 1000 base pairs [17]. This would also be a potent mechanism for toxicity. We therefore here report studies using NMR on the binding of CORM-3-derived Ru(II) to DNA, and show that it binds to guanosine N7 positions, leading eventually to precipitation of the DNA. Using an alkaline comet assay, which measures single-and double-stranded DNA breaks, we show that CORM-3 causes a marked increase in DNA strand breaks in RKO cells. We therefore conclude that Ru (as generated by dissolution of CORM-3 in water) binds to DNA and leads to DNA strand breaks. The binding is different from that seen with cisplatin, which may allow Ru to be developed as an alternative anticancer treatment, for example in cases of cisplatin toxicity or resistance, a topic of considerable current interest [24].

CORM-3 solutions bind to guanosine N7 in DNA
Binding of CORM-3-derived Ru(II) to DNA was assessed by NMR using a synthetic self-complementary B-DNA fragment, d(GTATGGCCATAC) 2 . This sequence was chosen because it contains a GpG sequence, which would allow 1,2-intrastrand binding, the most common binding site for the widely used anticancer drug cisplatin, Pt(NH 3 ) 2 Cl 2 [25]. It also contains a d(GpC) sequence, permitting 1,2-interstrand binding, also a common cisplatin binding mode.
The 1 H NMR spectrum of the DNA sequence was assigned using standard homonuclear methods (Table 1 and Figure 1). The 31 P spectrum was also assigned using 31 P-1 H COSY (Table 1, Figure 2). On addition of 1 equivalent of CORM-3, initial changes in the NMR spectrum were small, but after incubation for 3 days at 37°C , some large intensity changes were observable ( Figure 3). The assignments of these signals were checked by 2D NMR (Figure 4). The sample was then left in the dark at 37°C for a further month, after which time, further changes could be seen in the NMR spectrum. These further changes were in most cases a completion of the changes seen at 3 days, although a number of lower intensity broad signals grew in intensity, indicating secondary reaction sites with some crosslinking of DNA molecules. Most of the spectral changes were monotonic with time, though a few appeared and then disappeared later, for example a small signal seen at 8.13 ppm at 3 h but not subsequently. There is thus evidence for equilibration of Ru between different sites on the DNA, with the resultant pattern being presumably the thermodynamic equilibrium.
The NMR data are best summarised by mapping out the NOE connectivities that are affected significantly by incubation with CORM-3 ( Figure 5). The overwhelming majority of NOEs missing or greatly reduced are those involving guanosine H8 protons, implicating binding of Ru(II) to the adjacent N7 positions, which is the  An expansion of the region containing crosspeaks between H1 0 (vertical axis) and H6/8 (horizontal axis) protons. 1.5 mM duplex, mixing time 60 ms. The lines show the sequential NOE walk in rainbow colours used to assign the spectrum, starting in red at the 5 0 end. expected binding site for transition metals to B-DNA [26,27], including that of Ru(II) complexes [28,29]. The signal most affected, and already dramatically weaker at 3 h, is the H8 signal from G1 (Figure 3), which is clearly the most reactive site. This is at the end of our synthetic nucleotide, and is thus considerably more sterically available than any of the others. It is worth noting that the signal for this proton in the Ru complex is the only new signal that we were able to assign unambiguously in the complex, at 8.46 ppm. It has few NOEs to sugar protons in the NOESY spectrum (Figure 4), implying that attachment of Ru leads to weakening of the    interstrand hydrogen bonding pattern. The downfield shift of the G1 H8 signal is consistent with the binding of a strongly electron-withdrawing group such as Ru 2+ at the adjacent N7 position [30]. The guanosine residue least affected by addition of one equivalent of Ru is G6 (Figure 5), which suffers only a small drop in intensity. This was a surprising observation, because with cisplatin this residue would be the most strongly affected, since it can make both 1,2 intrastrand and 1,2 interstrand crosslinks. This implies that Ru is not involved in intramolecular crosslinks to any significant extent, an observation previously made for monofunctional ruthenium(II) arene complexes [28,31]. In this behaviour, CORM-3 appears to behave differently from the related cisand trans-[Ru(II)(DMSO) 4 Cl 2 ], both of which have been reported to crosslink two adjacent N7 positions [32,33].
The 31 P signal in DNA is sensitive to the double helix geometry, and changes markedly on either kinking or intercalation [34]. Neither was seen here, merely a loss in intensity of some 31 P signals, mainly those adjacent to guanosines (data not shown). This confirms the 1 H NMR result, implying structural weakening induced by Ru binding at isolated guanosine N7, but no crosslinking, kinking or intercalation of DNA.
Addition of a large excess of CORM-3 led to the appearance of a large number of additional signals, but the most marked effect was a gradual reduction in signal intensity throughout the spectrum, and a marked broadening of peaks, implying crosslinking and aggregation of DNA ( Figure 6). The NMR data therefore imply that Ru(II) binds at the most sterically accessible guanosine N7 sites, and does not make intramolecular crossbridges. At high concentration, it can gradually make intermolecular crossbridges, crosslinking different DNA double helices together, leading to broadening and loss of NMR signals.

CORM-3 causes DNA damage in RKO cells
We have previously shown that incubation of E. coli cells with CORM-2 leads to binding of Ru to DNA, with a stoichiometry of ∼3 per 1000 base pairs, after 1 h incubation of cells with 30 mM CORM-2 [17]. This should be enough to cause significant DNA damage. We therefore carried out experiments to test whether incubation of CORM-3 with mammalian cells leads to observable DNA damage. We used an alkaline comet assay. In this experiment, cells are grown, trypsinised and then incubated in the desired conditions. They are subsequently embedded in agarose, lysed, and electrophoresed in an alkaline buffer, followed by staining and visualisation. The alkaline conditions denature the DNA [35] and allow both single and double-strand breaks to be seen, as faster moving comet tails in the stained gel. Initial experiments showed that concentrations of CORM-3 greater than ∼30 mM are too toxic to cells and gave unreliable results, so we used 20 mM CORM-3 and a short incubation time. Our previous results showed that incubation of mammalian cells in serum-containing media protected against DNA damage, because the amino acids in the serum bind to CORM-3 and prevent its accumulation into cells. We therefore grew RKO cells (derived from a human rectal carcinoma) in their preferred RPMI medium, trypsinised, and resuspended in medium. They were then diluted into either phosphate buffered saline (PBS) or into serum-containing medium and incubated with 20 mM CORM-3 for 30 min, after which they were embedded in an agarose gel, lysed, and electrophoresed in alkaline conditions. The results are shown in Figure 7, and show a significant increase in DNA damage in cells incubated in PBS, but no effect in cells incubated in serum-containing medium, which contains amino acids and thus prevents accumulation of CORM-3 into the cells. We conclude that CORM-3 leads to DNA damage over the relatively short timeframe of this experiment, which is less than 30 min.

Discussion
We have previously shown that CORM-3 and CORM-2 kill cells not because of their released CO but because of the Ru(II) ion that they contain [16,17]. The ruthenium is concentrated inside cells, and binds to intracellular thiols, as found for example in glutathione and in cysteine residues on intracellular proteins. The results presented here confirm the cytotoxicity of Ru in CORM-3, and suggest that DNA damage is likely a major cause of the toxicity seen. In agreement with this finding, it has previously been shown that incubation of CORM-2 with E. coli leads to induction of the DNA repair gene recA [36], although Tavares et al. attributed this to an indirect response to the generation of reactive oxygen species (ROS). We note that the majority of studies on CORMs have seen a reduction in ROS, rather than an increase, following exposure to CORMs, but the data are unclear [3]. In light of our finding that very little CO is released from CORM-3 in typical growth conditions [16], a more likely explanation for the induction of recA is Ru-derived DNA damage. CORM-3 and CORM-2 are useful sources of Ru(II) [3]. The ligands maintain ruthenium in a Ru 2+ oxidation state. When dissolved in water, some of the ligands are replaced by water or by buffer, which makes the central Ru more accessible. This allows the CORMs to be taken up by cells, by a mechanism that is still not clear but may involve metal uptake systems. Ruthenium itself has no known natural biological function [3], so it has been suggested that it may be taken up using Fe 2+ or Cu 2+ uptake systems. However, it has also been suggested that the dipeptide permease/heme transporter DppABC may be involved [22]. Once inside the cell, the ruthenium reacts with intracellular nucleophiles, mainly thiols such as cysteine and glutathione. This alters the equilibrium of CORMs across the cell membrane and allows the ruthenium to attain much higher concentrations inside the cell than in the medium [16,17,37]. It would appear that the Ru(II) is sequestered inside the cell by thiol ligands, forming a reservoir of Ru(II), from which the Ru can be extracted under physiological conditions, for example to allow it to bind to DNA [38]. In [38] the bound thiol was readily oxidised, allowing it to be displaced by guanosine. However in general it is not clear to what extent thiol-bound intracellular Ru is available, for example to bind to DNA. The data presented here show that DNA binding is a slow and thermodynamically driven process, in comparison with thiol binding which is primarily kinetically driven. The implication is that the ruthenium transfers slowly (over a period of days) from thiols to DNA, although we have been unable to provide any clear proof that this occurs in our system.
The NMR results presented here demonstrate that the Ru(II) ion from CORM-3 binds to sterically accessible guanosine N7 positions, which appears to be generally the preferred binding site for Ru(II) [29,39]. The reaction is slow, taking several days to reach equilibrium at 37°C. We note that a similarly slow reactivity has been observed for other ruthenium complexes [40]. This is the same position that binds to cisplatin [26,27]. Our results however show a clear difference in that whereas cisplatin prefers intramolecularly bridged geometries, Ru(II) from CORM-3 clearly does not. The cytotoxicity of cisplatin is generally ascribed to its forming intramolecular crossbridges between pairs of purine N7, which leads to bending of DNA. Ruthenium does not do this, although it does form intermolecular crossbridges. Moreover, whereas platinum has affinity for adenosine [25], there is no indication from our results that ruthenium does so (see also [29]). The two metals therefore have a different molecular mechanism, which may provide a useful selectivity in both action and resistance [31]. In this context, it is worth noting that there have been several reports of Ru(II) complexes showing toxicity towards cisplatin-resistant cells [33,[41][42][43], including for the related complex RuCl 2 (DMSO) 4 [44]. They also have quite different kinetics, both for binding DNA and for entry to the cell. The aquated complexes formed from CORM-3 seem to behave differently from other aquated Ru(II) complexes [32,33]. It is thus clear that the specific ligand chemistry of a Ru(II) complex is critical for its biological effects.
Ruthenium compounds have been investigated as cancer therapeutics for decades, with general agreement that they show chronic rather than acute cytotoxicity, and that the nature of the complexation of the ruthenium ion outside the cell is critical for activity [45][46][47][48]. However, there has so far been little molecular understanding of the mode of action. We suggest that the insights from this study may be useful in refining research questions.
The slow binding equilibrium implies that on a cellular timescale, the cytotoxicity caused by CORM binding to DNA is unlikely to be reversible. It might therefore be expected to show extensive DNA damage, even if the amount of bound Ru is only of the order of one ion per 1000 base pairs or less. The results of the comet assay ( Figure 7) show that extensive DNA damage does indeed occur, and (as expected) is retarded by incubation of cells in complex RPMI medium. The DNA damage seen here occurs within 30 min; thus, although the reaction with DNA is slow, it is very effective at causing DNA damage.
In this paper and recent work [16,17], we have shown that CORM-3 and CORM-2 have two significant biological effects: they sequester a high proportion of intracellular thiols, and they also induce damage in DNA. Both of these are likely to be harmful to cells. This may provide a novel mechanism for cell death in the context of cancer therapeutics, compared for example to cisplatin, which works entirely by DNA binding. The existence of a large body of work on effects of CORMs on cells (reviewed in [3,49]) provides a good basis to move forward rapidly.

Materials
The DNA oligonucleotide was synthesised by Eurofins Genomics (Germany). It was dissolved to 5 mM in 30 mM potassium phosphate pH 7.8, and annealed by heating to 85°C and slowly cooling. The sample was lyophilised and redissolved in 100% D 2 O. TSP (0.1 mM) was added as reference. CORM-3 was prepared as a stock solution of 5-20 mM in 30 mM deuterated potassium phosphate buffer pH 7.8. Solutions were stored at 4°C in the dark and used on the day of preparation.

NMR experiments
NMR experiments were carried out on a Bruker Avance I (800 MHz, 1 H) and Bruker Avance DRX (202 MHz, 31 P) at 298 K. All homonuclear 2D experiments had 400 complex increments, with relaxation delays of 1.5 s. The mixing time for NOESY was 60-200 ms, and the spin lock time for TOCSY was 60 ms. The 31 P COSY had 64 complex increments with a 1 s relaxation delay. Assignments were made using a combination of COSY, TOCSY and NOESY, with sequential NOE walks for 1 0 and 2 0 /2 00 . Samples were left (at pH 7.8) at 37°C in the dark between NMR experiments and were monitored to check for lack of bacterial growth.
Cell culture and comet assays RKO cells were maintained as monolayers in RPMI medium supplemented with 10% foetal calf serum, 1% L-glutamine and 1% penicillin/streptomycin (Sigma) in a humidified atmosphere of 5% CO 2 at 37°C. The comet assay followed Breslin et al. [50]. Confluent cells were washed with PBS, treated with 2 ml trypsin solution (37.5 ml PBS, 30 ml trypsin stock [1 mg/ml trypsin, 1 mM HCl, 20 mM CaCl 2 ] and 7.5 ml 0.5 mM EDTA), and incubated for 2 min. Cells were centrifuged at 1000 rpm for 5 min, and resuspended in RPMI or PBS to a concentration of 3 × 10 5 cells ml −1 . An amount of 1 ml aliquots were taken, with half being treated with 20 mM CORM-3 and half being used as controls (30 min, 37°C). The cells were put on ice, centrifuged at 1000 rpm, and the pellet resuspended in 1 ml PBS. An amount of 150 ml of each sample was mixed with 150 ml 1.2% low-melting agarose at 42°C. An amount of 150 ml of this was pipetted onto a layer of set 0.6% normal agarose (150 ml) on a frosted glass cover slide, and a coverslip placed on top. The slides were left to set for 30 min at 4°C in the dark. The coverslips were then removed, and the slides were incubated in lysis buffer (2.5 M NaCl, 0.1 M EDTA, 10 mM Tris pH 10, 1% DMSO, 1% Triton X-100) at 4°C for 1 h in the dark. Slides were then placed in electrophoresis tanks in 1 mM EDTA, 50 mM NaOH, 1% DMSO and equilibrated for 45 m before running at 12 V for 25 m. Slides were neutralised by soaking overnight at 4°C in 0.4 M Tris pH 7.4, stained with SYBR green (Sigma) (1 : 20 000 dilution in PBS) for 5 m and scored using Comet Assay IV software. This measures the tail moment (tail length × % DNA in tail), a direct measure of the amount of DNA damage [51].

Data Availability
All data described here are available on reasonable application to the corresponding author (M.P.W.).


s2fieldsofstudy: Biology Medicine

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Endoplasmic reticulum stress contributed to inflammatory bowel disease by activating p38 MAPK pathway

Recent evidence suggests that endoplasmic reticulum (ER) stress plays a vital role in inflammatory bowel disease (IBD). Therefore, the aim of this study was to investigate the mechanism by which ER stress promotes inflammatory response in IBD. The expression of Gro-α, IL-8 and ER stress indicator Grp78 in colon tissues from patients with Crohn’s disease (CD) and colonic carcinoma was analyzed by immunohistochemistry staining. Colitis mouse model was established by the induction of trinitrobenzene sulphonic acid (TNBS), and the mice were treated with ER stress inhibitor tauroursodeoxycholic acid (TUDCA). Then the body weight, colon length and colon inflammation were evaluated, and Grp78 and Gro-α in colon tissues were detected by immunohistochemistry. Epithelial cells of colon cancer HCT116 cells were treated with tunicamycin to induce ER stress. Grp78 was detected by Western blot, and chemokines were measured by PCR and ELISA. The expression levels of Grp78, Gro-α and IL-8 were significantly upregulated in intestinal tissues of CD patients. Mice with TNBS induced colitis had increased expression of Grp78 and Gro-α in colonic epithelia. TUDCA reduced the severity of TNBS-induced colitis. In HCT116 cells, tunicamycin increased the expression of Grp78, Gro-α and IL-8 in a concentration-dependent manner. Furthermore, p38 MAPK inhibitor significantly inhibited the upregulation of Gro-α and IL-8 induced by tunicamycin. In conclusion, ER stress promotes inflammatory response in IBD, and the effects may be mediated by the activation of p38 MAPK signaling pathway.

Introduction
Inflammatory bowel disease (IBD) is a chronic gastrointestinal inflammatory disorder. Crohn's disease (CD) and ulcerative colitis (UC) are the two common types of IBD. Despite well-defined clinical manifestations, the pathogenesis of IBD has not been fully elucidated. It is generally accepted that gut microbiome dysbiosis, host immune response, environmental and genetic factors play key role in the initiation and progression of IBD. [1][2][3] Recent studies have shown that endoplasmic reticulum (ER) stress is associated with the susceptibility to IBD. 4 Various cellular stress conditions can result in the accumulation of unfolded or misfolded proteins in the ER lumen. The accumulation of unfolded and misfolded proteins in the ER lumen activates unfolded protein response (UPR) to repair protein folding defect and restore ER homeostasis. UPR signaling is mainly mediated by three protein sensors on the ER membrane: inositol-requiring transmembrane kinase/endonuclease 1 (IRE1), pancreatic ER eIF2α kinase (PERK), and activating transcription factor 6 (ATF6). Upon ER stress, activated IRE1a cleaves a 26-base intron at the dual stem loop structure of an mRNA that is then translated to produce Xbox-binding protein 1 (XBP1). XBP1 then induces the expression of genes that regulate protein folding, maturation, transport, phospholipid biosynthesis, and ER expansion to alleviate ER stress. Activated PERK phosphorylates α subunit of eukaryotic translation initiation factor 2 (eIF2α) on Ser51, which then suppresses translation and mitigates ER protein folding load. 5 The effect of ER stress on IBD is mainly mediated by inducing epithelial cell apoptosis, impairing the mucosal barrier function, regulating innate or adaptive immune response of the host cells, and triggering inflammatory signaling. 6,7 Chemokines are small heparin binding proteins that govern the migration of circulating leucocytes to the sites of inflammation. 8 Chemokines play a central role in the pathogenesis of both CD and UC, and their expression is consistently increased during the active phases of the diseases. In particular, interleukin 8 (IL-8) and its receptor are upregulated in intestinal mucosa. 9-12 IL-8 is one of the members of the CXC chemokine family which recruits neutrophil to inflamed sites. Previous studies have confirmed that IL-8 was expressed in the intestinal mucosa of IBD, and IL-8 expression level was correlated with disease activity. 13,14 Gro-α is another member of CXC family and promotes chemotaxis to neutrophil. Gro-α secretion in the serum increased in patients with UC and CD and then decreased after treatment. 15 Gro-α may be an important factor aggravating IBD inflammation, but its expression in IBD intestinal mucosal is still unknown. 16 Therefore, the aim of this study was to investigate the mechanism by which ER stress promotes inflammatory response in IBD focusing on the expression of chemokines.

Clinical samples
Colonic mucosal tissues were obtained from surgical specimens from 10 patients with CD. Control normal colonic mucosal specimens were obtained from pericarcinomatous tissue 8-10 cm away from the lesion location from 5 patients with colonic carcinoma. The surgical specimens were fixed with formalin, then dehydrated in alcohol, embedded in paraffin, sectioned, stained with hematoxylin-eosin, and mounted.

Animals
Six-week-old female BALB/c mice were purchased from Hunan Slake Jingda Laboratory Animal Co., Ltd (Changsha, China). The mice were acclimatized for 1 week before the experiment and housed individually in a room maintained at 22°C under a 12-h day/night cycle. All animal experiments were reviewed and approved by the Institutional Animal Care Committee of Changsha Central Hospital (Approval No. 20200512). Mouse IBD model was established by the induction with trinitrobenzene sulphonic acid (TNBS) following previous protocol. Briefly, 75 μL of TNBS (5% w/v) and 75 μL of ethanol absolute were fully mixed to form 130 μL of TNBS enema. After fasting for 24 h, the mice were anesthetized by intraperitoneal injection with 10% chloral hydrate (3 mL/kg). Next, Intragastric needle was slowly inserted into the intestinal cavity through the anus and TNBS enema was injected. The mice were inverted for about 5 min to ensure that the enema fluid was fully dispersed in the intestinal cavity. The model mice then received intraperitoneal injection of tauroursodeoxycholic acid (TUDCA) at dose of 50 mg/kg or normal saline as the control every day from the 2 nd day after enema. Disease activity index (DAI) score was calculated as the sum of the weight loss score, the diarrheal score and the hematochezia score following previous protocol. 17 At the end, all mice were killed by cervical dislocation.

Cell culture and treatment
Epithelial cells of colon cancer HCT116 were cultured in Dulbecco's modified Eagle's medium (DMEM) and high glucose supplemented with 10% fetal bovine serum (FBS),10,000 U/mL penicillin and 10,000 µg/mL streptomycin in a humidified incubator with 5% CO 2 at 37°C. Cells were treated with 5 µg/mL of tunicamycin (Cell Signaling, Danvers, MA, USA) for 24 h to induce ER stress. 18 Control cells were treated with 0.1% (v/v) solvent dimethyl sulfoxide (DMSO) as vehicle control.

Immunohistochemical staining
Paraffin-embedded colonic or ileal sections of patients and mice were cut at a 5 µm thickness. The sections were boiled in 10 mM sodium citrate buffer for antigen retrieval. After blocking endogenous peroxidase with 3% hydrogen peroxidase, sections were blocked for 30 min with 3% BSA, and then incubated with primary antibodies for Grp78 (Abcam, Cambridge, UK) (1:500), Gro-α (R&D Systems, Minneapolis, MN, USA) (1:4,000) and IL-8 antibody (R&D Systems) (1:25) for 24 h at 4°C. Phosphate buffered saline was used instead of primary antibodies as negative control. After incubation with horseradish peroxidase labeled secondary antibodies (Abcam) (1:200) at room temperature for 1 h, the sections were visualized with DAB, and counterstained with hematoxylin.

ELISA
The supernatants were collected from cells, and IL-8 levels in the supernatants were determined using Human IL-8/RANTES DuoSet kit (4A Biotech Co. Ltd., Beijing, China) following the manufacturer's instructions.

Statistical analysis
Statistical analyses were performed using GraphPad Prism 8.0, and all values were presented as means ±SD. Student's t-test were used to compare differences between groups, and p<0.05 was considered significant.

Upregulation of ER stress marker and chemokines in intestinal tissues of CD patients
To determine whether ER stress and chemokines are involved in CD, we performed immunohistochemistry staining for ER stress marker Grp78 and two chemokines IL-8 and Gro-α. Grp78, Gro-α and IL-8 were mainly localized to the epithelial lining of the gut and in Paneth cells, and were also localized in inflammatory cells.
Optical density analysis showed that the positive rates of Grp78, Gro-α and IL-8 staining in inflamed intestinal tissues of CD patients were higher compared with pericarcinomatous tissue (Figure 1 A-C).

TUDCA alleviated TNBS-induced colitis in mice
To confirm the involvement of ER stress and chemokines Groα and IL-8 in the development of IBD, we established mouse colitis model induced by TNBS.TNBS treated mice had significantly lower body weight, decreased colon length ( Figure 2A) and higher disease activity index score (DAI) than control mice ( Figure 2B). Histological analysis revealed that TNBS treated mice developed serious colitis ( Figure 2C). However, ER stress inhibitor TUDCA improved colon shortening in TNBS induced mice ( Figure 2A). Moreover, signs of colitis were markedly ameliorated, as evidenced by decreased DAI and histological score (Figure 2 B,C).

TUDCA inhibited the upregulation of Grp78 in colonic epithelia of mice
Furthermore, immunohistochemical staining of the colon tissues of the three groups of mice showed that the expression of Grp78 in TNBS model group was significantly higher than that of vehicle control group, but the expression of Grp78 in TNBS+TUDCA group was significantly lower than that of the TNBS model (Figure 3 A-D). These data indicated that TUDCA relieved ER stress in mouse colitis model.

Tunicamycin induced ER stress and increased Gro-α and IL-8 expression in HCT116 cells
To reveal the relationship of ER stress and chemokines in intestinal inflammation, human epithelial cells of colon cancer were cultured and treated with different concentrations of tunicamycin to induce ER stress. PCR analysis showed that tunicamycin increased the expression of Gro-α and IL-8 at mRNA levels in HCT116 cells (Figure 4 A,B). Western blot analysis of ER stress marker Grp78 confirmed that tunicamycin treated cells had higher expression of Grp78 compared to vehicle treated control cells ( Figure 4C). Furthermore, ELISA showed that IL-8 levels in the medium harvested from HCT116 cells treated with tunicamycin were higher compared to vehicle treated HCT116 cells ( Figure 4D). Collectively, these data demonstrated that tunicamycin induced ER stress and increased expression of chemokines Gro-α and IL-8 in vitro.

p38 MAPK signaling contributed to tunicamycininduced ER stress and increased expression of GRO-α and IL-8 in HCT116 cells
To identify signaling pathway responsible for the upregulation of Gro-α and IL-8 by tunicamycin, we used p38 MAPK kinase inhibitor SB203580. HCT116 were treated with different concentrations of SB203580 (5, 20,50 µM) in the presence or absence of tunicamycin, and supernatants were harvested 24 h after treatment. PCR analysis showed that SB203580 significantly decreased the  Article expression of Gro-α and IL-8 at mRNA levels in HCT116 cells treated with TM ( Figure 5 A,B). Western blot analysis of ER stress marker Grp78 confirmed that SB203580 inhibited TM induced expression of Grp78 compared to TM treated cells ( Figure 5C). Furthermore, ELISA showed that IL-8 levels in the medium harvested from HCT116 cells treated with SB203580 were significantly lower compared to TM treated HCT116 cells ( Figure 5D). Collectively, these data suggested that p38 MAPK signaling may mediate ER stress induced Gro-α and IL-8 expression in intestinal epithelial cells.

Discussion
In this study, we provide evidence that chemokines and ER stress played significant role in intestinal inflammation. A variety of chemokines have emerged as essential molecules in the pathogenesis of IBD. Multiple chemokines including Gro-α (CXCL1), ENA-78 (CXCL5), GCP2 (CXCL6) and IL-8 (CXCL8) are positively correlated with the inflammatory state in IBD patients. 18,19 IL-8, a classical pro-inflammatory chemokine, plays a role in the recruitment of neutrophils from the vasculature to the sites of infection or tissue injuries, and it has been shown that CXCL1 and CXCL8 were upregulated in colonic IBD. 20 By immunohistochemical staining we demonstrated that the expression of Grp78, Gro-α and IL-8 significantly increased in intestinal tissues of CD patients, compared with pericarcinomatous tissues.
Tunicamyin, a common inducer of ER stress, was previously shown to activate ER stress in intestinal epithelial cells. 21 In contrast, TUDCA can significantly reduce ER stress in colonic epithelial cells, and inhibit DSS-induced colon inflammation in mice. 22 Our results showed that the expression of Grp78 was upregulated by tunicamycin in a concentration dependent manner. In a mouse model of IBD induced by TNBS, inhibition of ER stress by TUDCA could reduce DAI score, increase survival rate, increase the colon length, maintain body weight, reduce the infiltration of inflammatory cells in the intestinal tissue, and alleviate TNBSinduced enteritis in mice. In addition, TUDCA inhibited the upregulation of Grp78 induced by tunicamycin in the colonic epithelia of mice.  Recent evidence suggests the role of MAPK signaling in the response to ER stress. For example, p38 was shown to promote ER stress induced apoptosis in fibroblasts and inflammation in macrophages. In addition, p38 can activate the transcription of CHOP through ATF6. 23 During inflammation, IL-17 enhanced TNF-α induced CXCL8 expression through p38 MAPK pathway, while p38 inhibitor SB203580 reversed the effect of IL-17 on CXCL8 mRNA, indicating that p38 MAPK signaling mediates the stabilization of CXCL8 mRNA by IL-17. 24 In this study, we found that SB203580 significantly inhibited the expression of ER stress marker Grp78 and chemokines Gro-α and IL-8 as well as the secretion of IL-8 in HCT116 cells treated with tunicamycin. Our findings suggested that p38 MAPK signaling may mediate ER stress induced inflammation in intestinal epithelial cells.
In conclusion, ER stress promotes inflammatory response in IBD, and the effects may be mediated by the activation of p38 MAPK signaling. Targeting p38 MAPK signaling may be a novel approach to the treatment of IBD. qRT-PCR analysis of IL-8 mRNA levels; GAPDH was set as an internal control; *p<0.05 vs vehicle. C) Western bot analysis of GRP78 protein levels. GAPDH was set as loading control. D) Supernatants were collected from HCT116 cells for ELISA to analyze the secretion of IL-8. Three independent experiments were performed in duplicate and data represented mean ± SD; *p<0.05 compared with TM treatment only group.


text:
The role of organ-deposited IgG in the pathogenesis of multi-organ and tissue damage in systemic lupus erythematosus

Systemic lupus erythematosus (SLE), often known simply as lupus, is a severe chronic autoimmune disease that is characterized by multi-organ and tissue damage and high levels of autoantibodies in serum. We have recently investigated, using animal models, the role of organ-deposited IgG autoantibodies in the pathogenesis of organ and tissue damage in SLE. We found that intra-organ injection of serum from mice with lupus (i.e., lupus mice) into healthy mice triggered inflammation in tissue and organs but that serum from other healthy mice did not, and that the severity of inflammation was related to the dose of serum injected. Immunohistochemistry showed that a large number of IgG molecules are deposited at the site of organ and tissue damage in lupus mice, and that IgG is a major contributor to the development of tissue inflammation triggered by serum from lupus mice or patients. The development of tissue inflammation induced by IgG in serum from lupus mice requires the presence of monocytes/macrophages, but not of lymphocytes or neutrophils; tumor necrosis factor (TNF)/tumor necrosis factor receptor 1 (TNFR1) and interleukin 1 (IL-1) also play essential roles in the development of tissue inflammation triggered by IgG. In addition, it has been found that TNFR1 inhibitors can suppress skin injury in lupus mice and that spleen tyrosine kinase (Syk) inhibitors, which can block the signaling transduction of IgG/Fc gamma receptors (FcγRs), can prevent and treat skin injury and kidney damage in lupus mice. We have also observed that lupus IgG might protect against bone erosion. Based on these results, we conclude that IgG plays a crucial role in the development of organ and tissue damage in SLE and in protecting bone erosion and arthritis, and we suggest that the IgG/FcγR signaling pathway is an important therapeutic target in SLE.

Introduction
Systemic lupus erythematosus (SLE), which affects mostly women of childbearing age, is a chronic severe autoimmune disease characterized by damage to multiple organs and tissues and a high level of autoantibodies in serum (1)(2)(3). The incidence and prevalence of SLE vary considerably between different regions of the world (1,4). In adults, the prevalence of SLE worldwide ranges from 30 to 150 per 100,000, and the incidence ranges from 2.2 to 23.1 per 100,000 per year (4). The typical initial symptoms of lupus are fever, erythema, and arthritis (5). Renal involvement is the most common manifestation: the prevalence of nephritis among lupus patients is 29-82% (6). Skin injury is the second most common symptom, and occurs in 70-85% of all lupus patients (7). Arthralgia is also common, and lupus patients with arthralgia often suffer from varying degrees of synovitis, but without bone erosion (8). Only 2.8-4.3% of patients have Jaccoud's arthropathy, which may result in joint deformities (8). With advancements in therapies, 5-, 10-, 15-, and 20-year survival rates have risen to 95%, 91%, 85%, and 78%, respectively (4). The main causes of death in SLE patients are infection, cardiovascular events, and active disease (4,9). Glucocorticoids and antimalarial agents (especially hydroxychloroquine) are the most important and most common first-line agents for the treatment of SLE, but B-celltargeting drugs have shown promising results (4,5). Belimumab, a humanized monoclonal antibody targeting soluble B-cell activating factor (BAFF), a B-cell stimulator, can significantly and sustainedly reduce the level of IgG autoantibodies, including anti-double-stranded DNA (anti-dsDNA), anti-Smith (anti-Sm), anti-cardiolipin, and anti-ribosomal P autoantibodies (10). In addition, the immature dendritic cell (iDC) vaccine, an underdeveloped therapy, has been shown to have a protective effect in lupus-like nephritis induced by pathogenic autoantibodies (11). These indicates that SLE IgG is involved in the therapy of SLE.
The organ and tissue damage that occurs in the region of IgG deposition, is due to infiltration by inflammatory cells, leading to the destruction of tissue organization. Severe damage to multiple organs is one of the leading causes of death in patients with SLE (3,12). Commonly damaged organs include the kidneys, skin, joints, liver, spleen, lungs, and brain (1-3).
The immune system is damaged in SLE patients, and B cells are abnormally over activated and produce multiple types of autoantibodies. Patients with SLE produce IgA, IgE, IgG, and IgM autoantibodies (12), but IgG autoantibodies predominate. Inflammation is the pathological basis of organ and tissue damage caused by the local deposition of IgG and immune complexes (ICs) (12)(13)(14)(15)(16)(17). The pathogenesis of SLE is closely associated with a high level of autoreactive IgG (2,3,18,19). However, the mechanism by which IgG causes organ and tissue damage is still unclear.
Tissue-deposited ICs formed by autoantibodies and autoantigens activate immune cells to produce inflammatory cytokines by binding to Fc gamma receptors (FcgRs) (20-24). Recently, we have demonstrated that organ-deposited IgG induces local inflammation by binding to FcgRs on the surface of monocytes/macrophages, resulting in multi-organ and tissue damage (20-28).
In this review, we summarize recent studies of the role of organdeposited IgG in the pathogenesis of inflammation and organ and tissue damage in SLE, and potential therapeutic targets in the IgG/ FcgRs signaling pathways, including studies investigating animal models, immune cells, cytokines ( Figure 1).

IgG deposited in tissues and organs
IgG and ICs have been found in the kidneys of SLE patients, and the most common autoantibodies identified are anti-dsDNA antibodies (18). IgM and IgG autoantibodies are deposited at the dermis-epidermis junction (29), forming so-called "lupus bands," which can be detected using direct immunofluorescence (30, 31). Some autoantibodies, including anti-galectin 3 antibody, antiacidic ribosomal protein P0 antibody (anti-RPLP0), and anti-Ro52 antibody, have been reported to be deposited in the skin and to be involved in the development of skin damage (32-34). We have demonstrated that IgG is deposited in the skin, liver, spleen, and joints of MRL/lpr mice (17, 23,27,28), and shown that the level of IgG deposited is correlated with the severity of tissue damage (17, 23,27,28). These findings suggest that tissuedeposited IgG may have an essential role in the pathogenesis of multi-organ and tissue damage in SLE.

Serum-induced inflammation in organs and tissue
We investigated whether or not serum containing a high level of IgG induces tissue inflammation in order to determine the role of tissue-deposited IgG in the multi-organ and tissue damage that occurs in SLE. Through intra-organ injection of serum from lupus mice and SLE patients (lupus serum), we have established an animal model of SLE organ and tissue damage (17, 20-22, 24, 27, 28). We found that intra-organ injection of serum from SLE patients and lupus mice, but not from healthy humans and mice, induced tissue inflammation in the skin, liver, spleen, and joints, and that the severity of the inflammation was dose-dependent (17, 23,27,28). Using toll-like receptor 4 (TLR4)-deficient mice that were lipopolysaccharide (LPS) resistant, we established that organ and tissue inflammation could be induced by serum from SLE patients and lupus-prone mice, but not by LPS contamination (17,27). We found that tissue inflammation caused by SLE serum appeared in the skin, liver, spleen, and joints a few hours after intra-organ injection, reached its peak after 3 days, and lasted for up to 14 days (17, 23,27,28).
Inflammatory skin reactions were induced by injecting serum from SLE patients into six different mouse strains: C57BL/6, SWISS, BALB/c, C3H/HeN, C3H/HeJ, and B-17 mice (23). Skin inflammation was induced by injection of serum only if the patients from which the serum was derived also exhibited skin inflammation and the severity of skin inflammation was not correlated with the severity of systemic disease (23); lupus serum had a synergistic effect with CpG DNA in inducing skin inflammation (23,35).
Other studies have also shown that intracerebroventricular injection of lupus serum induces an inflammatory response (36)(37)(38). These studies suggest that lupus serum can cause organ and tissue inflammation.

IgG plays a key role in inflammation induced by lupus serum
SLE is characterized by a high level of autoantibodies, particularly IgG. Many studies have shown that IgG is involved in the pathogenesis of organ and tissue inflammation and organ damage (17, 20-22, 24, 27, 28, 39, 40). We removed IgG from lupus serum to evaluate the role of IgG in the development of tissue inflammation. We found that the inflammation was significantly less severe in mice that received intra-organ injection of IgGdepleted lupus serum than in those injected with IgG-containing lupus serum (17, 23,27,28). The skin, liver, spleen, and joints could be directly inflamed by IgG isolated from lupus serum (17, 23,27,28). Furthermore, IgG extracted from lupus serum caused more severe inflammation than the same dose of IgG extracted from healthy serum (17, 23,27,28). The severity of inflammation was also related to the dose of lupus IgG (17, 23,27,28).
Recently, antibody glycosylation in autoimmune diseases has attracted attention, and the importance of Fc regions for inflammation development has been demonstrated (41,42). In addition to the antigen-binding site, the level of antibody glycosylation may differ in serum from healthy individuals and SLE patients. In vitro experiments have revealed that microglia are activated by phagocytosis of IgG (36). In a study in which mice received intrasplenic injection of lupus serum or of IgGdepleted lupus serum, the numbers of germinal centers(GCs) and IgG-secreting plasmacytes incresased after 8 days in the whole lupus serum group. (28). IgG has also been shown to induce monocyte differentiation into dendritic cells (DCs) (24). The role of IgG in organ and tissue damage in systemic lupus erythematosus (SLE). Lupus IgG and immune complexes (ICs) deposited in several organs and tissues cause multi-organ and tissue damage. IgG and ICs binding to Fc gamma receptors (FcgRs) activate monocytes/macrophages to secret cytokines. IgG inhibits osteoclastogenesis induced by the receptor activator of nuclear factor kappa B ligand (RANKL) to protect against bone erosion by binding to FcgRI and induces inflammation by activating the spleen tyrosine kinase (Syk)/nuclear factor kappa B (NF-kB) signal pathway to upregulate the transcription of inflammatory genes in monocytes/macrophages.
In addition, intrahepatic injection of IgG has also been shown to increase levels of ALT alanine transaminase (ALT) and aspartate aminotransferase (AST) in serum and to cause accumulation of apoptotic hepatocytes around the region of inflammation (17).
In another study, the severity of IgG-induced skin inflammation was found not to be related to the titer of the anti-dsDNA antibody and anti-Ro antibody (23). In addition, it has been reported that injection of anti-DNA antibodies causes proteinuria and promotes lupus nephritis in mice only when the antibodies were bound to the basement membrane (43). These findings demonstrate that IgG plays a key role in tissue inflammation induced by lupus serum.

FcgRI is important for the development of inflammation induced by SLE IgG
The effect of IgG is mediated by FcgR (44). FcgRI and FcgRIII are activating receptors, and FcgRII is the inhibitory receptor. The only inhibitory receptor, FcgRIIB, transmits its inhibitory signal through the immunoreceptor tyrosine-based inhibitory motif (ITIM) (45). Polymorphism of the genes encoding FcgRs has been reported and confirmed to be a heritable susceptibility factor for SLE (2,3,46). It has been reported that the FCGR2B-I232T genotype suppresses ligand binding, leading to low affinity for IgG (47). Because FcgRIIB is an inhibitory receptor, the low affinity for IgG contributes to a reduction in inhibitory signals, thus reducing inflammation, and alters the balance between inhibitory and activatory signals, which is associated with susceptibility to SLE (45,47).
Similarly, polymorphism of the genes encoding activating receptors also influences susceptibility to SLE (2,3,47). FcgRI is the only high-affinity FcgR for IgG, and is usually expressed on the surface of monocytes/macrophages (48). The immunoreceptor tyrosine-based activation motif (ITAM) embedded in the intracellular structure of FcgR is responsible for signal transduction of IgG and ICs (44, 48). The expression of FcgRI, but not of FcgRII and FcgRIII, has been found to be increased in monocytes from SLE patients, and the level of FcgRI expression was found to be related to the SLE disease activity index (SLEDAI) (49).
We used FcgRI-deficient mice with IgG-induced inflammation to investigate the role of FcgRI in the inflammation induced by SLE (49). The inflammation induced by IgG was reduced in FcgRI-deficient mice compared with wildtype mice (49). In addition, the expression level of FcgRI was associated with IgG deposition and skin inflammation in MRL/ lpr mice (37). Activation of the inflammatory signal pathway in monocytes/macrophages required FcgR expression (49).
However, in vitro experiments have shown that IgG reduces the surface expression of FcgRI, but not of FcgRII and FcgRIII, on monocytes/macrophages (27). N-glycosylation of antibodies and IgG subclasses affects the binding affinity of antibodies to different Fc receptors (50). Abnormal glycosylation of autoreactive IgG may have a higher affinity to FcgRI, suggesting that the molecular modification of autoreactive IgG may influence its pathogenicity. Further work is required to identify the Fc region provided by which state of the SLE IgG.

Monocytes/macrophages are required for inflammation induced by SLE IgG
Using various strains of cell-deficient and cell-depleted mice, we investigated the role of different types of immune cells in inflammation induced by IgG. Immunohistochemistry demonstrated that regions of tissue inflammation in the skin, kidneys, liver, and spleen are characterized by infiltration of a large number of monocytes/macrophages (17, 23,27,28,51). For example, F4/80 + macrophages were identified in the red pulp of the spleen (28), liver macrophages (Kupffer cells) were seen in areas of hepatitis induced by SLE IgG (17), and large numbers of CD11b + monocytes/macrophage and CD11c + DCs were detected in sites of skin inflammation; however, CD3 + T cells and CD20 + B cells were absent (23,52). The injection of lupus serum and IgG significantly reduced the inflammation caused by the depletion of monocytes/macrophages (17, 23,27,28).
Furthermore, skin inflammation induced by lupus IgG did not develop in Csf1-deficient mice lacking mature monocytes (21). Rag 1-deficient mice (which lack mature T and B cells but whose monocytes/macrophages are intact) were used to identify the role of T and B cells in IgG-induced inflammation (17,24,27). The lack of mature T and B cells did not affect the development of organ tissue inflammation induced by IgG (17,24,27). Neutrophil depletion by anti-Ly6G antibody injection did not affect the development of IgG-induced arthritis and dermatitis (20, 27). Another study showed that IgG induced lupus nephritis by activating macrophages (51).
In yet another study, IgG activated microglia to produce proinflammatory cytokines (53). In addition, polymorphism of the ITGAM (integrin subunit alpha M) gene (a gene associated with the activation of monocytes/macrophages) is associated with SLE susceptibility (11), and inflammation and organ damage in lupus-prone mice were suppressed, and splenic macrophages decreased, after treatment with the agonist of ITGAM (54). These findings indicated that monocytes/ macrophages but not T cells, B cells, or neutrophils, play a crucial role in IgG-induced inflammation.

TNF-a/TNFR1 plays a major role in the development of inflammation induced by SLE IgG
Many cytokines are involved in inflammation induced by IgG. Tumor necrosis factor alpha (TNF-a) is an important proinflammatory cytokine that is mainly secreted by macrophages and associated with SLE activity (55,56). TNF-a has been found to be highly expressed in the kidneys and skin of SLE patients (57), and in the joints, skin, and kidneys of MRL/lpr mice (21,27,58). In vitro stimulation of macrophages with IgG has been found to cause IC to produce TNF-a (17, 28, 51),, and the severity of inflammation induced by IgG in skin, liver, spleen, and joints has been shown to be significantly reduced in mice with a TNF-a deficiency (17,22,27,28). TNF exerts its effect through its receptors, such as TNFR1 and TNFR2. The skin inflammation induced by IgG was found to be reduced in mice with a TNFR1 deficiency, but not in those mice with a TNFR2 deficiency (24). Furthermore, we found that a deficiency of interleukin (IL) 1 (IL1) relieved inflammation induced by IgG by reducing the production of TNF-a (22). These results suggest that TNF-a/TNFR1 is crucial for inflammation induced by IgG in lupus serum. In addition, BAFF, produced by activated macrophages, was found in the region of inflammation to prolong the survival of macrophages in lupus nephritis (59,60). In addition, IL-10, interferon alpha (IFN-a), and interferon gamma (IFN-g) have been shown to increase CD64 expression in monocytes/macrophages (49,61). In one study, a lack of macrophage migration inhibitory factor (MIF) attenuated infiltration of macrophages and alleviated lupus nephritis (62).

Inhibitor of/FcgRs signaling pathway suppressed organ and tissue damage in lupus mice
Based on the findings from animal models of tissue inflammation induced by lupus IgG, the IgG/FcgRs signaling pathway is an important therapeutic target. Thus, we investigated whether an IgG/FcgRs signaling molecule inhibitor prevents and alleviates multi-organ and tissue damage in SLE. Spleen tyrosine kinase (Syk), a non-receptor tyrosine kinase of the Src family, plays a fundamental role in IgG/FcgRs signaling pathway conduction in autoimmune diseases (27,(63)(64)(65). FcgRI was required to activate Syk induced by SLE serum in macrophages (49). Syk inhibitors were used to treat lupus MRL/lpr mice, and have been found to prevent and treat damaged skin, kidneys, and spleen in MRL/lpr mice (28,63). In addition, Syk inhibitors reduced inflammation and damage in the spleen, liver, and skin induced by injection of IgG from lupus serum (17, 23,28). Intravenous immunoglobin (IVIG) is an anti-inflammation therapy used to treat various acute and chronic autoimmune diseases (66). Its anti-inflammation effect is due to the blocking of Fc receptors, which cannot bind with pathological autoantibodies (66). Glycosylation of the Fc region of IgG affects its function of inflammation modulation. The sialic acid-rich IgG fraction of IVIG has an improved anti-inflammatory activity (66). These results suggest that IgG/FcgRs signaling molecules may be a potential therapeutic target in multi-organ tissue damage of SLE.

IgG protects against bone erosion in SLE arthritis
Bone erosion is an important feature in inflammatory arthritis, such as rheumatoid arthritis; however, it does not occur in SLE arthritis. The receptor activator of nuclear factor kappa B ligand (RANKL) induces monocytes/macrophages to separate into osteoclasts that mitigate bone erosion (26). FcgRI is a costimulatory molecule of RANK, the receptor for RANKL that activates macrophage differentiation into osteoclasts (14). We investigated the role of IgG in macrophage differentiation to understand the mechanism by which IgG from lupus serum induces arthritis without bone erosion (27). We found that IgG inhibited osteoclastogenesis induced by RANKL in a dosedependent manner, and deficiency of FcgRII and FcgRIII did not influence the inhibitory effect of IgG in the formation of osteoclasts; both IgG from lupus serum and RANKL reduced the level of FcgRI (27). The inhibitory effect on osteoclastogenesis was greater at high doses than in at doses of IgG, and increasing doses of RANKL gradually blocked the inhibitory effects of lupus IgG. The stronger inhibitory effects of lupus IgG on RANKL-mediated osteoclastogenesis were presented in cells pre-treated for 24 hours when compared with cells treated with both RANKL and lupus IgG at the same time; at 24 hours after RANKL stimulation, the inhibitory effect of lupus IgG on osteoclastogenesis was eliminated (27). Based on these data, it appears that the relationship between IgG and RANKL is competition for FcgRI. IgG inhibits RANKL-induced osteoclastogenesis through competition for FcgRI binding. The binding of IgG to FcgRI may lead to a functional deficiency of FcgRI on the cellular membrane, which is required for RANKL-induced osteoclastogenesis. The competitive occupation of FcgRI by IgG may be exploited to develop therapeutic approaches to prevent bone destruction in autoimmune/inflammatory arthritis.

Conclusion
Based on these results, we conclude that IgG plays a crucial role in developing organ and tissue damage in SLE and in protecting against bone erosion in SLE arthritis, and that the IgG/FcgRs signaling pathway is an important therapeutic target in SLE.

Author contributions
Conception and design, G-MD; manuscript writing, WQ and TY; critical revision of the manuscript, G-MD; and, final approval of manuscript, all authors.

Funding
This study was supported by the National Natural Science Foundation of China (NSFC) (G-MD, 82171786).


s2fieldsofstudy:
Biology
Environmental Science

text:
Evolutionary History of DMSP Lyase-Like Genes in Animals and Their Possible Involvement in Evolution of the Scleractinian Coral Genus, Acropora

Dimethlysulfoniopropionate (DMSP) lyase is an enzyme that mediates cleavage of DMSP into dimethyl sulfide (DMS) and acrylate. DMS is an aerosol substance that may affect cloud formation, solar radiation and ocean temperatures. DMSP lyases in marine organisms, such as marine bacteria, release DMS, which might contribute to atmosphere-ocean feedback. Although DMSP lyases were first identified in marine bacteria, eukaryotic DMSP lyases or genes similar to DMSP lyase, DMSP lyase-like (DL-L) genes have been found not only in coccolithophores (Emiliania huxleyi) and symbiotic algae of the Family Symbiodiniaceae, but also in animals, including scleractinian corals (Cnidaria: Anthozoa: Hexacorallia). Comparative genomic analysis showed that gene expansion events of DL-L genes have occurred specifically in the scleractinian genus, Acropora. In the present study, we performed molecular identification of DL-L genes in Acropora digitifera. Thirteen full-length Open Reading Frames were isolated, confirming that these duplicated DL-L genes are likely expressed. A comprehensive survey of available transcriptomic databases revealed that DL-L genes have been identified not only in scleractinians (Hexacorallia), but also Octocorallia (Anthozoa) and even in a jellyfish (Cnidaria: Hydrozoa). Molecular phylogenetic analyses showed that although some sequences from cnidarian transcriptomic databases apparently originated with their symbiotic algae, cnidarian sequences from Anthozoa and Hydrozoa clustered together, indicating that these evolved from a gene in the last common ancestor of Cnidaria, dating to the Precambrian. Interestingly, cnidarian species possessing DL-L genes apparently occur only in coral reefs or shallow, warmer environments, suggesting that these genes may be essential for animals to survive in such environments. Acropora-specific duplicated DL-L genes, which originated during the past warm geological periods, may enable them to adapt to environmental changes.

INTRODUCTION
Coral reefs in tropical and subtropical waters harbor about 30% of all marine life, making them the most biodiverse habitats in marine ecosystems (Knowlton et al., 2010). In addition, coral reefs provide many fishery resources, protect coastlines, and fix carbon and nitrogen. Scleractinian or stony corals (Anthozoa, Cnidaria) are the main builders of coral reefs, forming massive calcium carbonate skeletons. Corals are metaorganisms or "holobionts" associated with a variety of microorganisms, such as dinoflagellates of the family Symbiodiniaceae and diverse communities of bacteria, archaea, fungi, viruses, and protists (Rohwer et al., 2002;Ainsworth et al., 2010). They rely mainly on symbiotic algae (Symbiodiniaceae) to trap solar energy for survival, growth, and calcification (Goreau and Goreau, 1959;Muscatine, 1990;Davy et al., 2012). In stony corals, dinoflagellate symbionts use the substrates (nitrogen and phosphorus) produced by their hosts for photosynthesis (Muscatine and Porter, 1977). Prokaryotic microorganisms provide sources of nitrogen, sulfur, and other elements to corals or release them into the water column (Ainsworth et al., 2010;Rädecker et al., 2015). These allow coral reefs to remain highly productive in oligotrophic waters, while also facilitating global biogeochemical cycles.
The sulfur cycle is one of the most important of these cycles, providing biosynthetic proteins and cofactors. Dimethylsulfoniopropionate (DMSP) is an organic sulfur compound that is abundant in marine surface waters. The majority of it is produced by phytoplankton and algae, but it has also been reported in some angiosperms and bacteria (Keller et al., 1989;Hanson et al., 1994;Paquet et al., 1994;Gage et al., 1997;Kocsis et al., 1998;Lyon et al., 2011;Kettles et al., 2014;Curson et al., 2017). Although functions of DMSP are not entirely understood, several physiological functions, such as osmoregulation, oxidative stress protection, and cryoprotection, have been demonstrated in phytoplankton and green algae (Kirst et al., 1991;Karsten et al., 1996;Sunda et al., 2002;Lesser, 2006;Husband et al., 2012;Curson et al., 2018). Biological functions such as chemoattraction, predator deterrence, and mediation of bacterial virulence have been identified (Wolfe and Steinke, 1996;Miller and Belas, 2004;Barak-Gavish et al., 2018). The most notable and important aspect of DMSP is that it is a precursor of dimethyl sulfide (DMS) (van Boeckel and Stefels, 1993), which is a major source of sulfur in the atmosphere, promoting cloud formation and combating greenhouse gasses. DMSP is cleaved by an enzyme called DMSP lyase (EC 4.4.1.3), which converts DMSP to DMS. Recently, studies reported that DMSP lyase products, DMS and acrylate, may have ecological roles such as chemical defenses against predation (Strom et al., 2003;Teng et al., 2021) and enhancing predation by grazers (Shemi et al., 2021). When DMS enters the atmosphere, it is oxidized into aerosol particles that can induce cloud formation and increase reflectivity. This may reduce light levels and water temperatures in marine environments, thus contributing to local climate regulation ( Figure 1) (Ayers and Gras, 1991;Vallina and Simo, 2007), although some studies suggest that sea-air DMS flux is low, and that the role of DMS in climate regulation is likely insignificant (Woodhouse et al., 2010;Quinn and Bates, 2011).
Marine phytoplankton and macroalgae are considered the primary producers of DMSP, while marine bacteria are the primary degraders (Stefels et al., 2007;Reisch et al., 2011). DMSP is released from marine phytoplankton and macroalgae upon cellular lysis caused by zooplankton grazing, viral infection, and senescence ( Figure 1) (van Boekel and Stefels, 1993;Wolfe et al., 1997;Hill et al., 1998). Bacteria acquire DMSP mainly from the ocean and demethylate it into methanethiol and acetaldehyde, turning DMSP into DMS via the lysis pathway ( Figure 1). DMSP lyase was first identified in bacteria, and at least eight prokaryotic DMSP lyase genes have been identified to date .
Some studies have reported that eukaryotic phytoplankton, including dinoflagellates and algae, possess DMSP lyase activity (Stefels et al., 1995;Steinke and Kirst, 1996;Steinke et al., 1998;Niki et al., 2000;Yoch, 2002), but the responsible gene was difficult to isolate. In 2015, a gene, Alma l, was identified from the eukaryotic coccolithophore, Emiliania huxleyi, and was shown to have high DMSP lyase activity (Alcolombri et al., 2015). It has been suggested that Alma 1 can directly lyse algal DMSP, releasing acrylate and DMS ( Figure 1). Interestingly, sequences similar to Alma 1, DMSP lyase-like (DL-L) genes were also found in scleractinian corals and Symbiodiniaceae (Alcolombri et al., 2015). Furthermore, recent comparative genomic analysis of scleractinian and cnidarian genomes showed that numbers of DL-L genes in genomes of the scleractinian coral genus, Acropora, are significantly larger than in other cnidarian genomes and that gene expansion events specifically occurred in Acropora (Shinzato et al., 2021a).
Acropora is currently the most studied coral genus in the world (Maor-Landaw and Levy, 2016), as it includes the most widespread, abundant, and diverse scleractinian corals on Earth (Wallace and Rosen, 2006). Approximately 180 species have been recorded from the Red Sea to the Indo-Pacific Ocean and the Caribbean Ocean (Wallace, 1999;Veron, 2000;van Oppen et al., 2001). With the increase of research data, more genetic databases (genome and transcriptome databases) and molecular techniques (CRISPR/Cas9 and gene knockdown) have been developed (Yasuoka et al., 2016;Cleves et al., 2018;Ying et al., 2019;Shinzato et al., 2021a). Genetic analyses suggest that Acropora has been evolutionarily very successful and may have become a dominant genus through acquisition and expansion of gene families (Shinzato et al., 2011;Shinzato et al., 2021a). Examples of Acropora-specific gene duplication and expansion events have been reported (Hislop et al., 2005;Shinzato et al., 2021a). Symbiotic relationship with Symbiodiniaceae may also have contributed to rapid adaptation to changing environments (Qin et al., 2019). Interestingly, Shinzato et al. (2021a) showed that DL-L genes are the most diversified gene family in Acropora genomes and that possible coral-or cnidarian-specific stress response genes, including Caspase-X and SCRiPs, have also been tandemly duplicated in Acropora genomes. In this study, to better understand evolution of DL-L genes in animals and their possible involvement in Acropora evolution, we first confirmed that DL-L genes are indeed expressed in A. digitifera as possible functional mRNAs without in-frame stop codons. Then, we performed a comprehensive database survey and molecular phylogenetic analyses to investigate the existence of DL-L genes in various eukaryotes, including scleractinian corals, cnidarians, and other animals, to gain insight into the evolutionary origin of DL-L genes in the animal kingdom.

MATERIALS AND METHODS
Cloning of DL-L Genes From Acropora digitifera The same RNA samples used in previous studies including different developmental stages of A. digitifera (eggs, blastulae, gastrulae, planula larvae, early polyps, and adult branches) were used for cDNA synthesis (Shinzato et al., 2011;Shinzato et al., 2021a;Yoshioka et al., 2021). Samples were homogenized in TRIzol reagent (Invitrogen) on ice, and total RNA was extracted following the manufacturer's instruction and Chiu et al. (2020). Extracted total RNA was treated with recombinant DNase I (Roche) to remove contaminating genomic DNA. First-strand cDNA was synthesized from 5 mg of DNase-treated RNA using SuperScriptTM III reserve transcriptase (Thermo Fisher Scientific). Possible DL-L genes of A. digitifera were derived from predicted genes of A. digitifera (Shinzato et al., 2021a), and primers amplifying full length open reading frames (ORFs) were designed for each gene (Supplementary Table 1). PCR products were cloned into the pGEM-T Easy cloning vector (Promega), transformed into competent cells (Escherichia coli, DH5a strain), and plasmids were sequenced at a commercial laboratory (Eurofin Genomics, Tokyo, Japan).

Identification of DL-L Genes From Publicly Available Eukaryote and Cnidarian Transcriptomic Databases
Amino acid sequences were deduced from cloned cDNA sequences using the ExPASy translate tool (https://web.expasy.org/translate/; Gasteiger et al., 2003). To search for genes similar to DMSP lyase gene from transcriptomic databases, the deduced amino acid sequences of cloned A. digitifera DL-L genes, Emiliania huxleyi Alma1 (KR703620.1), and Symbiodinium A1 DMSP lyase (P0DN22) were used as query sequences in homology searches against the Transcriptome Shotgun Assembly (TSA) database at the National Center for Biotechnology Information (NCBI) using tBlastn (A. digitifera DL-L genes: e-value cutoff 1e -50 , query coverage >80%, Emiliania huxleyi Alma1 and Symbiodinium A1 DMSP lyase: e-value cutoff 1e -5 , query coverage >70%). Sequences satisfying these criteria were retrieved, and amino acid sequences were prepared using the ExPASy translate tool.

DL-L Genes Isolated From Acropora digitifera RNAs and Their Gene Expression Patterns
Among 18 DL-L genes predicted from the A. digitifera genome (Shinzato at el., 2021a), we successfully obtained 13 DL-L gene sequences with full-length mRNA coding regions from cDNA ( Table 2) as shown by previous studies (Shinzato et al., 2021a;Shinzato et al., 2021b;Yoshioka et al., 2021). Gene expression of five genes was significantly changed (one upregulated and four downregulated) by increased sea water temperature (25°C to 30°C, q < 0.05, Supplementary Table 2).

DL-L Genes in Eukaryotes
Among the 6,258 eukaryotic TSA databases deposited in NCBI, we identified 123 gene sequences from 72 species that showed similarity to A. digitifera DL-L genes and satisfied the above criteria (tBlastn e-value cutoff 1e-50 , query coverage >80%). Most sequences were identified from the Cnidaria (stony corals, sea anemones, jellyfish and hydra), phytoplankton (Phyla Haptophyta, Myzozoa, Ochrophyta) and macroalgae (Phyla Chlorophyta and Rhodophyta) ( Table 1). Unexpectedly, four sequences were also identified from other animals, including a ctenophore (Beroe forskalii), a mollusk (Limacina retroversa) and arthropods (Dendroctonus frontalis and Pleuromamma xiphias) ( Table 1). We further investigated the amino acid sequence similarity of these sequences with the only known DMSP lyases from eukaryotes to date (Alcolombri et al., 2015), Emiliania huxleyi Alma1 and Symbiodinium A1 DMSP lyase (Supplementary Table 3). Most eukaryote sequences similar to A. digitifera DL-L genes ( Table 1) also showed sequence similarities to both DMSP lyases, but percent identities were lower than those to A. digitifera DL-L genes (Supplementary Table 3).
In order to infer evolutionary relationships of the identified sequences from eukaryotes and the four sequences from noncnidarian animals, we performed molecular phylogenetic analyses with amino acid sequences of the 13 A. digitifera DL-L genes as representatives of Cnidaria and bacterial DMSP lyase genes from prokaryotes as an outgroup. Sequences from prokaryotes and eukaryotes separated into two distinct clusters, indicating that all of the sequences obtained from eukaryote TSA databases originated from eukaryotes, and were not contaminants from symbiotic bacteria (Figure 3; Supplementary Figure 2). All sequences from A. digitifera clustered, demonstrating that they originated from the host coral ( Figure 3; Supplementary Figure 2). None of the sequences from non-cnidarian animals clustered with those from A. digitifera. Clustering of the D. frontalis sequence with dinoflagellate sequences displayed 100% bootstrap support, indicating that this sequence was derived from symbiotic or adhesive algae, not from the animal itself. However, the other three sequences from B. forskalii, P. xiphias and L. retroversa did not cluster with those of A. digitifera and other eukaryotes (Figure 3; Supplementary Figure 2); thus, the evolutionary origin (animals or other eukaryotes) are unclear.

DL-L Genes in Cnidaria
Among 137 Cnidarian TSA databases in NCBI, we found sequences similar to DL-L genes from 35 species ( Table 2). Most of these sequences were from Acropora millepora and A. tenuis and had been previously identified (Alcolombri et al., 2015;Shinzato et al., 2021a). Interestingly, possible DL-L genes were found not only in the subclass Hexacorallia, including stony corals, sea anemones, and zoanthids, but also in the Octocorallia, including soft corals and blue corals, and even from Hydroidolina, including jellyfishes and fire corals ( Table 2). We also investigated amino acid sequence similarity of these cnidarian sequences with known eukaryote DMSP lyases from E. huxleyi and Symbiodinium A1 (see above), and all sequences except one from Seriatopora caliendrum had significant similarities with these eukaryote DMSP lyases (Supplementary Table 4).
Coral and/or cnidarian transcriptomic databases often contain sequences originating with symbiotic algae, as well as associated microorganisms. In order to identify which sequences FIGURE 2 | Molecular characterization of DL-L genes in Acropora digitifera. Multiple sequence alignment of deduced amino acid sequences from 13 DL-L genes of A. digitifera and Emiliania huxleyi, Alma1 (KR703620.1). Identical residues are indicated in the same color. Lengths of amino acid sequences are shown at the right and positions of the Asp/Glu/hydantoin racemase superfamily conserved domain are shown in light yellow and the two putative active site residues are shown in red boxes. were from animal hosts, we further performed molecular phylogenetic analyses of sequences from cnidarian TSA databases together with DL-L genes from the Symbiodiniaceae, coccolithophores and prokaryotes ( Figure 4A). Six distinct clades were supported by strong bootstrap probabilities (931 00%). Most of the sequences belonging to the Hexacorallia clustered together in Clade 1 with A. digitifera DL-L genes, indicating that these originated from hexacorallian hosts.  Interestingly, Clavularia sp. of the Octocorallia and Velella velella of the Hydroidolina were also relegated to Clade 1, indicating that these share a common ancestral gene with hexacorallian species. The family Symbiodiniaceae is genetically diverse, and includes multiple genera (LaJeunesse et al., 2018). As expected, some Cnidarian sequences clustered with sequences of symbiotic algae (Symbiodiniaceae) and were assigned to Clades 3-5 ( Figure Table 2). Different cnidarian species harbor different types of symbiotic algae; thus, Clades 3-5, containing Symbiodiniaceae could reflect different types of symbiotic algae. All DL-L genes of the coccolithophore E. huxleyi within the Haptophyta clustered in Clade 2, and bacterial DMSP lyase genes clustered in Clade 6. Taken together, all DL-L genes of Cnidaria, including scleractinian corals, soft corals, and jellyfishes, evolved from an ancestral gene that already existed in the last common ancestor of Anthozoa and Hydrozoa, and have completely different evolutionary backgrounds from those of coccolithophores, the Family Symbiodiniaceae, or prokaryotes.

DL-L Genes
It has been reported that coral reefs are hotspots for DMSP, which have been attributed to symbiotic algae of corals (Broadbent et al., 2002;Broadbent and Jones, 2004;Broadbent and Jones, 2006;Swan et al., 2012). Recent studies have shown that adult and juvenile A. tenuis and A. millepora without symbiotic algae both produce DMSP (Raina et al., 2013). DL-L genes have also been discovered, not only in Acropora genomes (Alcolombri et al., 2015;Shinzato et al., 2021a), but also in genomes of Montipora, Astreopora, Goniastrea, and two corallimorpharians, Amplexidiscus and Discosoma (Shinzato et al., 2021a).
In this study, we successfully identified full-length ORFs of 13 DL-L genes, which are expressed in A. digitifera, and we identified DL-L genes from a variety of scleractinian lineages (Figures 1, 4A; Supplementary Figure 3). The earliest coral fossil record of a Scleractinian dates to the middle Triassic (240 Ma) (Stanley, 2003). Most extant scleractinians are classified into two major clades, known as the Complexa (complex corals) and Robusta (robust corals), and are assumed to have diverged in the Late Carboniferous (300 Ma) (Romano and Palumbi, 1996;Romano and Cairns, 2000). In genomes of robust corals, a DL-L gene has been detected only in Goniastrea aspera to date (Shinzato et al., 2021a). However, in this study, we identified animal-type DL-L genes from four robust corals, including Cyphastrea serailia, Favites colemani, Favia lizardensis, and Seriatopora caliendrum ( Figure 4B; Table 2), indicating that DL-L genes have been preserved in a variety of complex and robust corals, but were lost from some lineages.

Ancient Origin of DL-L Genes in Cnidarians
Although a large proportion of DL-L genes were identified in corals and cnidarians, we also identified similar sequences from non-cnidarian animals, B. forskalii, P. xiphias and L. retroversa. However, none of these share a common ancestry with coral DL-L genes (Figure 3). Although the origins of these sequences are not clear, we suggest that they may have come from symbiotic/ adhesive algae, as in the case of a sequence from D. frontalis (Figure 3). Based on our phylogenetic analyses using currently available transcriptomic databases, we conclude that, to date, only cnidarians possess DL-L genes that are clearly of animal origin. Further addition of more phytoplankton DL-L genes may reveal the origins of these from non-cnidarian species.
The Phylum Cnidaria contains three clades (subphyla): Anthozoa (comprising the Octocorallia, Hexacorallia, and Ceriantharia), Endocnidozoa (a clade of parasites) and Medusozoa (consisting of the Cubozoa, Hydrozoa, Scyphozoa, and Staurozoa) (Collins, 2009;Kayal et al., 2018). In this study, we identified eukaryotic DL-L genes, not only in the Scleractinia and Corallimorpharia but also in the Zoantharia (Hexacorallia), Alcyonacea (Octocorallia), and Hydrozoa (Anthoathecata, Hydroidolina) ( Figure 4B; Table 2). This indicates that the last common ancestor of Anthozoa and Hydrozoa possessed a DL-L gene and that this gene has been inherited in a wide range of cnidarian species. The earliest cnidarian fossils occur in strata of the Ediacaran (560 Ma, Liu et al., 2014), and together with molecular and paleontological analyses, suggest that the phylum Cnidaria probably originated during the Precambrian Eon from the Cryogenian to the Ediacaran (700-595 Ma, Peterson et al., 2004;Erwin et al., 2011). Divergence of 2 major taxa (Anthozoaria and Medusozoa) may have occurred before the Cambrian (543 Ma) (Cartwright et al., 2007;Park et al., 2012). Consequently, we hypothesize that the DL-L gene may be have originated from an ancient gene that already existed in the last common ancestor of Cnidaria in the pre-Cambrian ( Figure 4B).
How did cnidarians, including corals, obtain DL-L genes? Shinzato et al. (2021a) hypothesized that DL-L genes may have been acquired by the common ancestor of scleractinians and corallimorpharians, both of which are hexacorallians (Anthozoa), via horizontal gene transfer from symbiotic Symbiodiniaceae or Emiliania. However, as mentioned above, a variety of anthozoans and hydrozoans possess these genes ( Figure 4A). McFadden et al. (2021) suggested that the Anthozoa probably arose in the Cryogenian to Tonian periods (648-894 Ma) and lacked photosymbionts. In the Devonian (383 Ma) from the Palaeozoic Era, anthozoans of the Scleractinia first formed associations with photosymbionts, followed by alcyonacean octocorals (318 Ma) and corallimorpharians (312 Ma), and photosymbioses have been gained and lost repeatedly in all orders through the Jurassic (199-151 Ma, McFadden et al., 2021). In addition, the fossil record shows that scleractinian corals had photosymbionts in most of the upper Triassic since the Mesozoic Era (Stolarski et al., 2011;Frankowiak et al., 2016). Recent molecular dating estimates suggest that the earliest diversification of the Symbiodiniaceae occurred in the Jurassic (∼160 Ma, LaJeunesse et al., 2018) and the first E. huxleyi appeared 270,000 years ago (Thierstein et al., 1977;Paasche, 2001), which were much later than the origin of the Anthozoa. Our phylogenetic analysis (Figure 4) confirms distinct ancestries of DL-L genes in Symbiodiniaceae and cnidarians, indicating that they have completely different evolutionary backgrounds. Therefore, acquisition of DL-L genes via horizontal gene transfer from symbiotic algae (Shinzato et al., 2021a) may not have occurred. Based on the present findings, we propose an updated hypothesis: DL-L genes in cnidarians are "ancient genes" in the animal kingdom, dating back to the pre-Cambrian.
DL-L Genes: Essential for Survival in Coral Reef or Shallow and Warm Water Environments?
As of 10 th February 2022, 137 transcriptomic databases from 91 cnidarian species had been registered in NCBI TSA.
Interestingly, we realized that all cnidarian species, except V. velella, possessing animal type DL-L genes (15 species) are limited to coral reefs (Table 1). V. velella (Figure 4; Table 2), known as "by-the-wind sailor", is a cosmopolitan, free-floating, colonial hydrozoan that lives mainly at the water/air interface and floats in temperate and tropical seas (Araya and Aliaga, 2018). This implies that DL-L genes may be essential for animals to survive in coral reef or warm, shallow-water environments, although we acknowledge that publicly available transcriptomic databases of cnidarians to date may be limited to coral reef species.
C o m p a r a t i v e g e n o m i c a n a l y s i s u s i n g r e p o r t e d scleractinian coral genomes showed that DL-L genes are the most diversified gene family among gene families that significantly increased number of genes in the last common ancestor of Acropora (Shinzato et al., 2021a). Why did gene duplication events of DL-L gene specifically occur in the Acropora lineage? The earliest fossil records of Acropora are known from the late Paleocene (65-54 Ma) in Somalia and Austria (Carbone et al., 1993;Baron-Szabo, 2006), indicating that Acropora has existed for more than 50 million years. Acropora was the main reef builder in the Oligocene (28-23 Ma) of Greece and Early Miocene of Egypt (Schuster, 2002), and was widely distributed throughout the Miocene in different regions, including the Indo-Pacific and Caribbean (Budd, 2000). It is suggested that Acropora began to spread throughout the world in the Cenozoic and species diversification occurred in the Eocene and Oligocene (around 25-50 Ma). Molecular dating analysis using wholegenome data indicates that the Acropora ancestor survived warm periods without sea ice from the mid or late Cretaceous to the Early Eocene, when gene expansion of DL-L genes occurred specifically in the Acropora ancestor (Shinzato et al., 2021a). In addition, Acropora species also have high concentrations of DMSP compared to other corals (Broadbent et al., 2002;Guibert et al., 2020), suggesting that Acropora corals actively utilize DMSP and that high concentrations of DMSP may have triggered Acroporaspecific duplication of DL-L genes. Several studies have shown that DMSP is involved in a wide range of coral stress responses, including responses to heat, sunlight, air exposure, and hyposalinity (Sunda et al., 2002;Raina et al., 2009;Deschaseaux et al., 2014;Aguilar et al., 2017). Not all A. digitifera DL-L genes (5 out of 18) responded to increased sea water temperature and response patterns of differentially expressed genes vary (1 upregulated and 4 downregulated, Supplementary Table 2), suggesting that the functions of the DL-L genes in A. digitifera have also diverged. Taken together, diversified DL-L genes in Acropora may have acquired new or different functions, not only mediating cleavage of DMSP into DMS, but that they may assist Acropora in adapting to environmental changes, for example from intense heat, light, and salinity (Shinzato et al., 2021a). Eventually Acropora may become the dominant coral genus in extant coral reefs.
The functions of DL-L genes in corals are completely undetermined at this stage. Interestingly, although overexpression of the Alma l genes from E. huxleyi and Symbiodinium A1 in E. coli cells had high DMSP lyase activities, an Alma1 homolog from A. millepora showed almost no DMSP lyase activity (Alcolombri et al., 2015), indicating that not all DL-L genes in eukaryotes retain their DMSP lyase function, and that acquisition of other functions or functional differentiation may have occurred during the process of gene duplication. In particular, duplicated DL-L genes in Acropora will need to be investigated to determine which genes have DMSP lyase activity. We identified expressed sequences of DL-L genes from A. digitifera RNA, and these could be used for molecular and functional characterization in DMSP lyase assays. We also found that expression levels of DL-L genes were indeed diverse in A. digitifera. Therefore, even if some DL-L genes in corals do not function as DMSP lyases, they may have other functions enabling adaptation to coral reef or warm, shallowwater environments. Identifying their actual biological functions in Acropora corals will be important to understand not only adaptation mechanisms to shallow and warmer environments, but also the impact of Acropora corals on the sulfur cycle in the oceans.

DATA AVAILABILITY STATEMENT
The datasets presented in this study can be found in online repositories. The names of the repository/repositories and accession number(s) can be found in the article/ Supplementary Material.

AUTHOR CONTRIBUTIONS
CS conceptualized, commenced and supervised the project. Y-LC performed the lab work, database searches and molecular phylogenetic analyses. Y-LC and CS analyzed the data and wrote the manuscript. All authors contributed to the article and approved the submitted version.

SUPPLEMENTARY MATERIAL
The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmars.2022.889866/ full#supplementary-material Supplementary Figure 1 | Domain structure of the 13 deduced amino acid sequences of A. digitifera and Emiliania huxleyi, Alma1 (KR703620.1). Lengths of amino acid sequences are shown at the right and positions of the Asp/Glu/ hydantoin racemase superfamily conserved domain are shown in gray boxes. Scale bar=20 aa, aa: the number of amino acids.
Supplementary Figure 2 | Maximum likelihood analysis of DL-L genes identified from transcriptomic databases of eukaryotes. A total of 65 DL-L genes from prokaryotes and eukaryotes, including A. digitifera DL-L genes, were aligned using MAFFT v7.310. Then, 337 gap-trimmed aligned amino acid sequences were used for the phylogenetic analysis. Each species name is followed by a different symbol indicating the taxonomy of the species. The bar indicates 0.5 substitutions per site in aligned regions.
Supplementary Figure 3 | Maximum likelihood analysis of DL-L genes identified from transcriptomic databases of cnidarians. A total of 65 DL-L genes from cnidaria and prokaryotes including, A. digitifera DL-L genes, were aligned using MAFFT v7.310. Then, 337 gap-trimmed aligned amino acid sequences were used for the phylogenetic analysis. Each species name is followed by a different symbol indicating the taxonomy of the species. The bar indicates 0.6 substitutions per site in aligned regions.
Supplementary Table 1 | List of the primers for cloning PCR analysis.